The field of transgenics was initially developed to understand the action of a single gene in the context of the whole animal and the phenomena of gene activation, expression, and interaction. This technology has also been used to produce models for various diseases in humans and other animals and is amongst the most powerful tools available for the study of genetics, and the understanding of genetic mechanisms and function. From an economic perspective, the use of transgenic technology for the production of specific proteins or other substances of pharmaceutical interest (Gordon et al., 1987, Biotechnology 5: 1183-1187; Wilmut et al., 1990, Theriogenology 33: 113-123) offers significant advantages over more conventional methods of protein production by gene expression.
Heterologous nucleic acids have been engineered so that an expressed protein may be joined to a protein or peptide that will allow secretion of the transgenic expression product into milk or urine, from which the protein may then be recovered. These procedures have had limited success and may require lactating animals, with the attendant costs of maintaining individual animals or herds of large species, including cows, sheep, or goats.
The hen oviduct offers outstanding potential as a protein bioreactor because of the high levels of protein production, the promise of proper folding and post-translation modification of the target protein, the ease of product recovery, and the shorter developmental period of chickens compared to other potential animal species. The production of an avian egg begins with formation of a large yolk in the ovary of the hen. The unfertilized oocyte or ovum is positioned on top of the yolk sac. After ovulation, the ovum passes into the infundibulum of the oviduct where it is fertilized, if sperm are present, and then moves into the magnum of the oviduct, lined with tubular gland cells. These cells secrete the egg-white proteins, including ovalbumin, lysozyme, ovomucoid, conalbumin and ovomucin, into the lumen of the magnum where they are deposited onto the avian embryo and yolk.
2.1 Microinjection
Historically, transgenic animals have been produced almost exclusively by microinjection of the fertilized egg. Mammalian pronuclei from fertilized eggs are microinjected in vitro with foreign, i.e., xenogeneic or allogeneic, heterologous DNA or hybrid DNA molecules. The microinjected fertilized eggs are then transferred to the genital tract of a pseudopregnant female (e.g., Krimpenfort et al., in U.S. Pat. No. 5,175,384). However, the production of a transgenic avian using microinjection techniques is more difficult than the production of a transgenic mammal. In avians, the opaque yolk is positioned such that visualization of the pronucleus, or nucleus of a single-cell embryo, is impaired thus preventing efficient injection of the these structures with heterologous DNA. What is therefore needed is an efficient method of introducing a heterologous nucleic acid into a recipient avian embryonic cell.
Cytoplasmic DNA injection has previously been described for introduction of DNA directly into the germinal disk of a chick embryo by Sang and Perry, 1989, Mol. Reprod. Dev. 1: 98-106, Love et al., 1994, Biotechnology 12: 60-3, and Naito et al., 1994, Mol. Reprod. Dev. 37:167-171; incorporated herein by reference in their entireties. Sang and Perry described only episomal replication of the injected cloned DNA, while Love et al. suggested that the injected DNA becomes integrated into the cell's genome and Naito et al. showed no direct evidence of integration. In all these cases, the germinal disk was not visualized during microinjection, i.e., the DNA was injected “blind” into the germinal disk. Such prior efforts resulted in poor and unstable transgene integration. None of these methods were reported to result in expression of the transgene in eggs and the level of mosaicism in the one transgenic chicken reported to be obtained was one copy per 10 genome equivalents.
2.2 Retroviral Vectors
Other techniques have been used in efforts to create transgenic chickens expressing heterologous proteins in the oviduct. Previously, this has been attempted by microinjection of replication defective retroviral vectors near the blastoderm (PCT Publication WO 97/47739, entitled Vectors and Methods for Tissue Specific Synthesis of Protein in Eggs of Transgenic Hens, by MacArthur). Bosselman et al. in U.S. Pat. No. 5,162,215 also describes a method for introducing a replication-defective retroviral vector into a pluripotent stem cell of an unincubated chick embryo, and further describes chimeric chickens whose cells express a heterologous vector nucleic acid sequence. However, the percentage of G1 transgenic offspring (progeny from vector-positive male G0 birds) was low and varied between 1% and approximately 8%. Such retroviral vectors have other significant limitations, for example, only relatively small fragments of nucleic acid can be inserted into the vectors precluding, in most instances, the use of large portions of the regulatory regions and/or introns of a genomic locus which, as described herein, can be useful in obtaining significant levels of heterologous protein expression. Additionally, retroviral vectors are generally not appropriate for generating transgenics for the production of pharmaceuticals due to safety and regulatory issues.
2.3 Transfection of Male Germ Cells, Followed by Transfer to Recipient Testis
Other methods include in vitro stable transfection of male germ cells, followed by transfer to a recipient testis. PCT Publication WO 87/05325 discloses a method of transferring organic and/or inorganic material into sperm or egg cells by using liposomes. Bachiller et al. (1991, Mol. Reprod. Develop. 30: 194-200) used Lipofectin-based liposomes to transfer DNA into mice sperm, and provided evidence that the liposome transfected DNA was overwhelmingly contained within the sperm's nucleus although no transgenic mice could be produced by this technique. Nakanishi & Iritani (1993, Mol. Reprod. Develop. 36: 258-261) used Lipofectin-based liposomes to associate heterologous DNA with chicken sperm, which were in turn used to artificially inseminate hens. There was no evidence of genomic integration of the heterologous DNA either in the DNA-liposome treated sperm or in the resultant chicks.
Several methods exist for transferring DNA into sperm cells. For example, heterologous DNA may also be transferred into sperm cells by electroporation that creates temporary, short-lived pores in the cell membrane of living cells by exposing them to a sequence of brief electrical pulses of high field strength. The pores allow cells to take up heterologous material such as DNA, while only slightly compromising cell viability. Gagne et al. (1991, Mol. Reprod. Dev. 29: 6-15) disclosed the use of electroporation to introduce heterologous DNA into bovine sperm subsequently used to fertilize ova. However, there was no evidence of integration of the electroporated DNA either in the sperm nucleus or in the nucleus of the egg subsequent to fertilization by the sperm.
Another method for transferring DNA into sperm cells was initially developed for integrating heterologous DNA into yeasts and slime molds, and later adapted to sperm, is restriction enzyme mediated integration (REMI) (Shemesh et al., PCT International Publication WO 99/42569). REMI utilizes a linear DNA derived from a plasmid DNA by cutting that plasmid with a restriction enzyme that generates single-stranded cohesive ends. The linear, cohesive-ended DNA together with the restriction enzyme used to produce the cohesive ends is then introduced into the target cells by electroporation or liposome transfection. The restriction enzyme is then thought to cut the genomic DNA at sites that enable the heterologous DNA to integrate via its matching cohesive ends (Schiestl and Petes, 1991, Proc. Natl. Acad. Sci. USA 88: 7585-7589).
It is advantageous, before the implantation of the transgenic germ cells into a testis of a recipient male, to depopulate the testis of untransfected male germ cells. Depopulation of the testis has commonly been by exposing the whole animal to gamma irradiation by localized irradiation of the testis. Gamma radiation-induced spermatogonial degeneration is probably related to the process of apoptosis. (Hasegawa et al., 1998, Radiat. Res. 149:263-70). Alternatively, a composition containing an alkylating agent such as busulfan (MYLERAN™) can be used, as disclosed in Jiang F. X., 1998, Anat. Embryol. 198(1):53-61; Russell and Brinster, 1996, J. Androl. 17(6):615-27; Boujrad et al., Andrologia 27(4), 223-28 (1995); Linder et al., 1992, Reprod. Toxicol. 6(6):491-505; Kasuga and Takahashi, 1986, Endocrinol. Jpn 33(1):105-15. These methods likewise have not resulted in efficient transgenesis or heterologous protein production in avian eggs.
2.4 Nuclear Transfer
Nuclear transfer from cultured cell populations provides an alternative method of genetic modification, whereby donor cells may be sexed, optionally genetically modified, and then selected in culture before their use. The resultant transgenic animal originates from a single transgenic nucleus and mosaics are avoided. The genetic modification is easily transmitted to the offspring. Nuclear transfer from cultured somatic cells also provides a route for directed genetic manipulation of animal species, including the addition or “knock-in” of genes, and the removal or inactivation or “knock-out” of genes or their associated control sequences (Polejaeva et al., 2000, Theriogenology, 53: 117-26). Gene targeting techniques also promise the generation of transgenic animals in which specific genes coding for endogenous proteins have been replaced by exogenous genes such as those coding for human proteins.
The nuclei of donor cells are transferred to oocytes or zygotes and, once activated, result in a reconstructed embryo. After enucleation and introduction of donor genetic material, the reconstructed embryo is cultured to the morula or blastocyte stage, and transferred to a recipient animal, either in vitro or in vivo (Eyestone and Campbell, 1999, J Reprod Fertil Suppl. 54:489-97). Double nuclear transfer has also been reported in which an activated, previously transferred nucleus is removed from the host unfertilized egg and transferred again into an enucleated fertilized embryo.
The embryos are then transplanted into surrogate mothers and develop to term. In some mammalian species (mice, cattle and sheep) the reconstructed embryos can be grown in culture to the blastocyst stage before transfer to a recipient female. The total number of offspring produced from a single embryo, however, is limited by the number of available blastomeres (embryos at the 32-64 cell stage are the most widely used) and the efficiency of the nuclear transfer procedure. Cultured cells can also be frozen and stored indefinitely for future use.
Two types of recipient cells are commonly used in nuclear transfer procedures: oocytes arrested at the metaphase of the second meiotic division (MII) and which have a metaphase plate with the chromosomes arranged on the meiotic spindle, and pronuclear zygotes. Enucleated two-cell stage blastomeres of mice have also been used as recipients. In agricultural mammals, however, development does not always occur when pronuclear zygotes are used, and, therefore, MII-arrested oocytes are the preferred recipient cells.
Although gene targeting techniques combined with nuclear transfer hold tremendous promise for nutritional and medical applications, current approaches suffer from several limitations, including long generation times between the founder animal and production transgenic herds, and extensive husbandry and veterinary costs. It is therefore desirable to use a system where cultured somatic cells for nuclear transfer are more efficiently employed.
What is needed, therefore, is an efficient method of generating transgenic avians that express a heterologous protein encoded by a transgene, particularly in the oviduct for deposition into egg whites.